Embryo Culture: Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY™

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For further volumes: http://www.springer.com/series/7651

Embryo Culture Methods and Protocols Edited by

Gary D. Smith University of Michigan, Ann Arbor, MI, USA

Jason E. Swain University of Michigan, Ann Arbor, MI, USA

Thomas B. Pool Fertility Center of San Antonio, San Antonio, TX, US

Editors Gary D. Smith University of Michigan Ann Arbor, MI, USA

Jason E. Swain University of Michigan Ann Arbor, MI, USA

Thomas B. Pool Fertility Center of San Antonio San Antonio, TX, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) ISBN 978-1-61779-970-9 ISBN 978-1-61779-971-6 (eBook) DOI 10.1007/978-1-61779-971-6 Springer New York Heidelberg Dordrecht London Library of Congress Control Number: 2012941744 © Springer Science+Business Media, LLC 2012 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher’s location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is a brand of Springer Springer is a part of Springer Science+Business Media (www.springer.com)

Preface Assisted reproductive technologies have had a profound impact on biomedical research through transgenic animals, food supply and production, as well as genetic gain of domestic species, and treatment of human infertility. Understanding regulatory pathways of early mammalian embryo development, preimplantation embryo cellular differentiation, and pluripotency of the embryonic inner cell mass has been pivotal to advances in embryo culture, transgenic mouse production, and biomedical advances in basic research, understanding the pathophysiology and progression of diseases, and development of novel treatments and cures. In 2007, Drs. Capecchi, Evans, and Smithies were awarded the Nobel Prize in Physiology or Medicine for their groundbreaking discoveries in relation to utility of embryonic stem cells, DNA recombination, and refinement of gene targeting in mice. Central to transgenic animal technology is mammalian embryo culture. While assisted reproductive technologies for the treatment of infertility can be controversial and have a relatively brief lifespan, they have also touched many families’ lives in very positive ways. The pioneering utility of assisted reproductive technologies by Patrick Steptoe and Robert Edwards at Bourn Hall in Cambridge, England culminated in the birth of Louis Brown on July 25, 1978. This was not a chance happening. Efforts of numerous individuals, including Dr. Edwards, pre-1970s in studying and understanding the basic concepts of oocyte biology, sperm function, processes of fertilization and embryo production, culture, and selection paved the way for the clinical success of treating infertility with assisted reproductive technologies. These scientific and laboratory skills, in conjunction with enhanced understanding and regulation of the reproductive endocrine system and clinical advances in laparoscopy refined by Dr. Steptoe, led to the initial success in treating infertility. Human infertility afflicts approximately 10 % of the world’s population. Continued efforts of many basic, translational, and clinical scientists have improved the success of treating infertility with assisted reproductive technologies. Today it is estimated that more than four million children have been born by in vitro fertilization. As is fitting, Dr. Edwards was awarded the 2010 Nobel Prize in Physiology or Medicine for the development of in vitro fertilization therapies. Central to these therapies is mammalian embryo culture. While significant advances in embryo culture have occurred over the last few decades, there is still room for improvement. This book is targeted to basic science researchers, clinical laboratorians, and fertility healthcare providers. An emphasis is placed on advances, controversies, and opportunities to improve human embryo culture. With this said, important reviews regarding utility, strengths, and weaknesses of model systems in designing and testing culture alterations for improved human embryo culture are essential and discussed. Embryo culture media are dissected, and components discussed, by experts with emphasis on impact on cell function and embryo development. This is followed by debate of benefits and limitations of various embryo culture systems and environmental influence beyond the soluble component of embryo culture. Within most chapters practical laboratory protocols and/or specific procedures are presented. Finally, a discussion of emerging methodologies and impact of embryo culture on epigenetics and offspring health is presented to set the stage for future research and laboratory application involving embryo culture.

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As is true in most professional crafts, careers, and science, mentoring and collaboration are keys to passing on knowledge and facilitating discovery. Interaction between scientists and clinicians only hastens advancements in obtaining basic biology information and translating these data to improved clinical care. We are fortunate in the field of embryology and assisted reproductive technologies to have pioneers that truly had vision and recognized the pathway to success was through discovery, training, sharing knowledge, and mentoring the next generation of scientists. Many such mentors are key contributors to this collection of works. We would like to express our appreciation to the distinguished authors who have contributed to this project. Your efforts have made this a great mentoring tool. Finally, we thank Humana Press, Methods in Molecular Biology Series, and Dr. John M. Walker, series editor, for their support and confidence in our abilities to organize, institute, and complete this project. Ann Arbor, MI Ann Arbor, MI San Antonio, TX

Gary D. Smith Jason E. Swain Thomas B. Pool

Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 Historical Background of Gamete and Embryo Culture . . . . . . . . . . . . . . . . . . . . . Jacques Cohen and Don Rieger 2 Utility of Animal Models for Human Embryo Culture Development: Rodents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gary D. Smith 3 Utility of Animal Models for Human Embryo Culture Development: Domestic Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rebecca L. Krisher 4 Utility of Animal Models for Human Embryo Culture: Nonhuman Primates. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Calvin Simerly and Gerald Schatten 5 Media Composition: Salts and Osmolality . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jay M. Baltz 6 Media Composition: Energy Sources and Metabolism. . . . . . . . . . . . . . . . . . . . . . . Deirdre Zander-Fox and Michelle Lane 7 Media Composition: Amino Acids and Cellular Homeostasis . . . . . . . . . . . . . . . . . Franchesca D. Houghton 8 Media Composition: Macromolecules and Embryo Growth . . . . . . . . . . . . . . . . . . Marius Meintjes 9 Media Composition: Antioxidants/Chelators and Cellular Function . . . . . . . . . . . . Catherine M.H. Combelles and Margo L. Hennet 10 Media Composition: pH and Buffers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jason E. Swain 11 Media Composition: Growth Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aparna Hegde and Barry Behr 12 Culture Systems: Single Step. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ronit Machtinger and Catherine Racowsky 13 Culture Systems: Sequential . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Patrick Quinn 14 Culture Systems: Embryo Co-Culture. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yves J.R Ménézo, Edouard Servy, Anna Veiga, André Hazout, and Kay Elder 15 Culture Systems: Low-Oxygen Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Borut Kovačič 16 Culture Systems: Embryo Density. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael L. Reed

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17 Culture Systems: Air Quality. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 Theodore Thomas 18 Culture Systems: Mineral Oil Overlay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325 Dean E. Morbeck and Phoebe H. Leonard 19 Culture Systems: Physiological and Environmental Factors That Can Affect the Outcome of Human ART . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333 Don Rieger 20 Culture Systems: Fluid Dynamic Embryo Culture Systems (Microfluidics) . . . . . . . 355 André Monteiro da Rocha and Gary D. Smith 21 Human Embryo Culture Media Comparisons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 Thomas B. Pool, John Schoolfield, and David Han 22 Culture Systems: Embryo Culture and Monozygotic Twinning. . . . . . . . . . . . . . . . 387 Amy E. Sparks 23 Embryo Culture and Epigenetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 Brenna A. Market Velker, Michelle M. Denomme, and Mellissa R.W. Mann

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors JAY M. BALTZ • Department of Obstetrics and Gynecology, Ottawa Hospital Research Institute, University of Ottawa, Ottawa, ON, Canada BARRY BEHR • IVF Laboratory, Department of OB/GYN, Division of Reproductive Endocrinology and Infertility, Stanford University, Stanford, CA, USA JACQUES COHEN • Tyho-Galileo Research Laboratories & Reprogenetics, West Orange, NJ, USA CATHERINE M.H. COMBELLES • Biology Department, Middlebury College, McCardell Bicentennial Hall, Middlebury, VT, USA MICHELLE M. DENOMME • Departments of Obstetrics & Gynecology and Biochemistry, Children’s Health Research Institute, University of Western Ontario, London, ON, Canada KAY ELDER • Bourn Hall Clinic, Cambridge, UK DAVID HAN • Department of Statistics, University of Texas at San Antonio, San Antonio, TX, USA ANDRÉ HAZOUT • UNILABS France, Paris, France APARNA HEGDE • Department of OB/GYN, Division of Reproductive Endocrinology and Infertility, Stanford University, Stanford, CA, USA MARGO L. HENNET • Biology Department, Middlebury College, McCardell Bicentennial Hall, Middlebury, VT, USA FRANCHESCA D. HOUGHTON • Faculty of Medicine, University of Southampton, Southampton General Hospital, Southampton, UK PHOEBE H. LEONARD • Division of Reproductive Endocrinology and Infertility, Department of Obstetrics and Gynecology, Mayo Clinic, Rochester, MN, USA BORUT KOVAČIČ • Department of Human Reproduction and Gynecologic Endocrinology, University Medical Centre Maribor, Maribor, Slovenia REBECCA L. KRISHER • National Foundation for Fertility, Lone Tree, CO, USA MICHELLE LANE • Department of Obstetrics and Gynaecology, School of Pediatrics and Reproductive Health, University of Adelaide, Adelaide, SA, Australia RONIT MACHTINGER • Obstetrics, Gynecology & Reproductive Biology, Brigham and Women’s Hospital, Boston, MA, USA MELLISSA R.W. MANN • Departments of Obstetrics & Gynecology and Biochemistry, Children’s Health Research Institute, University of Western Ontario, London, ON, Canada MARIUS MEINTJES • Frisco Institute for Reproductive Medicine, Frisco, TX, USA YVES J.R. MÉNÉZO • UNILABS France, Paris, France DEAN E. MORBECK • Division of Reproductive Endocrinology and Infertility, Department of Obstetrics and Gynecology, Mayo Clinic, Rochester, MN, USA THOMAS B. POOL • Fertility Center of San Antonio, San Antonio, TX, USA PATRICK QUINN • Research and Development, Sage IVF, A CooperSurgical Company, Redmond, OR, USA ix

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CATHERINE RACOWSKY • Obstetrics, Gynecology & Reproductive Biology, Brigham and Women’s Hospital, Boston, MA, USA MICHAEL L. REED • Center for Reproductive Medicine of New Mexico, Albuquerque, NM, USA DON RIEGER • Research and Development, LifeGlobal, LLC, Guelph, ON, Canada ANDRÉ MONTEIRO DA ROCHA • Department of Cell and Developmental Biology, University of Michigan, Ann Arbor, MI, USA GERALD SCHATTEN • Division of Developmental & Regenerative Medicine, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA JOHN SCHOOLFIELD • Department of Periodontics, University of Texas Health Science Center at San Antonio, San Antonio, TX, USA EDOUARD SERVY • The Servy Institute, Augusta, GA, USA CALVIN SIMERLY • Pittsburgh Development Center of Magee-Women’s Research Institute, Departments of Obstetrics, Gynecology & Reproductive Sciences and Cell Biology & Physiology, University of Pittsburgh School of Medicine, Pittsburg, PA, USA GARY D. SMITH • University of Michigan, Ann Arbor, MI, USA AMY E. SPARKS • IVF and Reproductive Testing Laboratories, Department of Obstetrics and Gynecology, Center for Advanced Reproductive Care, University of Iowa Hospitals and Clinics, Iowa City, IA, USA JASON E. SWAIN • University of Michigan, Ann Arbor, MI, USA THEODORE THOMAS • Reproductive Technology Laboratory, Martha Jefferson Hospital, Charlottesville, VA, USA ANNA VEIGA • Servei de Medicina de la Reproducció, Institut Universitari Dexeus, Barcelona, Spain BRENNA A. MARKET VELKER • Departments of Obstetrics & Gynecology and Biochemistry, Children’s Health Research Institute, University of Western Ontario, London, ON, Canada DEIRDRE ZANDER-FOX • Department of Obstetrics and Gynaecology, School of Pediatrics and Reproductive Health, University of Adelaide, Adelaide, SA, Australia

Chapter 1 Historical Background of Gamete and Embryo Culture Jacques Cohen and Don Rieger Abstract The first tissue culture media were developed nearly 150 years ago by Ludwig and Ringer. These were simple salt solutions, which were initially based on the chemical properties of blood serum. The second generation of culture media was developed more than a century later, in the 1970s, aiming to mimic the reproductive environment. In the 1990s, simplex optimization was used to design the third group of media, to some extent ignoring existing formulations and principles. Simultaneous with the development of culture media, it became evident that it was necessary to carefully control the culture conditions, including temperature, pH, osmolarity, and air quality. Equally important was the development of instruments and tools specifically designed for cell tissue culture such as the inverted microscope, the incubator, the Petri dish, sterile plasticware, the laminar flow cabinet, and air filtration equipment. Key words: Embryo culture, Media, Temperature, pH, Osmolarity, Dishes, Air filtration, Historical aspects

1. Introduction The first tissue culture media were developed nearly 150 years ago by Ludwig and Ringer. These were simple salt solutions, which were initially based on the chemical properties of blood serum. Knowledge was gained during those early years about the biochemistry of metabolism in mammals and humans, particularly osmolarity, pH and temperature. This knowledge provided the basis for making many changes to the salt solutions that were already in use. The second generation of culture media was developed more than a century later, in the 1970s, aiming to mimic the reproductive environment (the so-called “back to nature” principle). The third group of media was designed to optimize growth in vitro, to some extent ignoring existing formulations and principles. To formulate these media, the performance of each ingredient was evaluated separately using a process referred to as “simplex optimization”—a system

Gary D. Smith et al. (eds.), Embryo Culture: Methods and Protocols, Methods in Molecular Biology, vol. 912, DOI 10.1007/978-1-61779-971-6_1, © Springer Science+Business Media, LLC 2012

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developed first for the mouse in the laboratory of John Biggers at Harvard University nearly 20 years ago. This work was supported by the United States National Institutes of Health (NIH) in part to define culture media for use in the human while to circumvent the moratorium on human embryo research, which has been in place since 1979. The US National Institute of Child Health and Human Development provided a large program grant bearing the name National Cooperative Program on Non-Human In Vitro Fertilization and Preimplantation Development; this was more colloquially known as “The Culture Club.” Simultaneous with the development of culture media for studying fertilization and embryogenesis, scientists and industrialists became more aware of the need to control environmental conditions and started developing or perfecting instruments specifically for cell tissue culture such as the inverted microscope, the incubator, the Petri dish, sterile plasticware, the laminar flow cabinet, and High Efficiency Particulate Air (HEPA) filtration. This chapter reviews historical components of gamete and embryo culture and in three sections: (1) development of culture media, (2) the study of physiological conditions, and (3) the development of common embryology-specific tools. 1.1. Development of Culture Media 1.1.1. Ringer’s Saline Solution and Its Derivatives

Sydney Ringer of University College in London, a British clinician and pharmacologist, developed a simple salt solution based on the constituents of blood serum for the study of the beating frog heart in vitro (1, 2). It was composed of sodium chloride (NaCl), potassium chloride (KCl), calcium chloride (CaCl2), and a low concentration of sodium bicarbonate (NaHCO3, 2.7 mM). Ringer’s was a modification of another solution used in the 1860s by Carl Ludwig who was trying to develop a perfusion model for organs. The accidental addition of London tap water by Ringer’s assistant was found to be crucial to maintain regular heartbeat and resulted in a marked improvement over previous perfusion solutions. The component in tap water that had been missing before was CaCl2. Ringer’s solution was modified by Locke and Rosenheim a quarter century later (3), notably by the addition of 11.1 mM glucose, for the study of the perfused rabbit heart. This solution is often referred to as Ringer–Locke’s. Maurice Tyrode, an American pharmacologist, added sodium phosphate (NaH2PO4) and magnesium chloride (MgCl2) and increased the concentration of NaHCO3 to 11.9 mM for use in the study of rabbit intestine (4). Krebs and Henseleit (5) removed the glucose, but increased the concentration of NaHCO3 to 25 mM, for the study of nitrogen metabolism in rat tissue. This concentration of NaHCO3 in Krebs–Ringers bicarbonate solution (KRB) is what is conventionally used in conjunction with 5 % CO2 in many tissue culture media in order to effect physiological pH.

1 1.1.2. Embryo Culture Media Based on Krebs– Ringers Bicarbonate Solution

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John Hammond was the first director of the new Animal Reproduction Unit of the Animal Research Station in Cambridge (UK) in 1949 (6). He was the first to observe mouse embryos cleaving in vitro and 8-cell embryos reaching the blastocyst stage. He was presumably also the first to observe the hatching process in vitro. He is considered the father of embryo culture for having set in motion a host of studies undertaken in the 1950s by experimental embryologists interested in obtaining fertilization and extended embryo culture in vitro, allowing continuous observations to be made on developmental stages from just after sperm penetration to immediately preimplantation. The general plan was to model preimplantation development in vitro, although some scientists also suggested continuing to grow the fetus in vitro. Hammond used a salt solution containing sodium chloride, potassium chloride, calcium chloride, magnesium chloride and some glucose for culture. To this he added about 5 % egg white. The embryos were flushed and placed in a serological tube, sealed, and cultured in air for 48 h. Only a few of the 8-cell embryos could develop further under those conditions. Hammond demonstrated that glucose was essential for compaction and development of the blastocyst. He was presumably the first to publish on the use of test tubes for mouse embryo culture. In 1956, Wesley Whitten from Australia showed that more than 90 % of 8-cell mouse embryos would develop to the blastocyst stage when cultured in KRB supplemented with 5.55 mM glucose, penicillin, streptomycin, and bovine serum albumin (7). McLaren and Biggers (8) were the first to show that such blastocysts would produce live young when transferred to recipient mothers. This was the first step on the road to in vitro fertilization (IVF). Whitten (9) added lactate to his 1956 formulation and was able to culture 2-cell mouse embryos from outbred animals to the blastocyst stage. However, he could not achieve the same with zygotes. This observation led to the discovery of the so-called “2-cell block” in mouse embryo culture. Both Hammond and Whitten used serological or test tubes with volumes of media 0.98 and the standard error of duplicates is 150 mM, remove the embryo from the micromanipulation dish. Tighter pipets may squeeze the embryo and cause the blastomeres to escape the zona pellucida. Gently wash the embryo through several drops of pre-equilibrated culture media before returning it to the culture drop for continued incubation (see Note 5). 3.1.2. Laser-Assisted Hatching of CleavageStage Embryos

1. Prepare the assisted hatching dish with 10 ml droplets of micromanipulation medium for embryos that are to be hatched. Quickly cover the drops with washed mineral or paraffin oil, making sure the drops are completely covered with oil. Warm the dish on a 37 °C surface for at least 15 min prior to micromanipulation (see Note 6). 2. Load the embryo(s) into the micromanipulation medium drops. 3. Focus on the embryo on high power (40×) and orient the embryo to hatch at a gap between blastomeres or over an area of fragmentation. If performing blastomere biopsy, position the embryo so the desired blastomere can be accessed from the area you intend to hatch. 4. With the embryo in place, switch to the laser objective. 5. Focusing on the outer edge of the zona pellucida, fire the laser. Moving inward continue to fire until you have cut a 20–25 mM hole though the zona pellucida (see Note 7). 6. Using a pipet with an inner diameter of >150 mM, remove the embryo from the micromanipulation dish. Tighter pipets may squeeze the embryo and cause the blastomeres to escape the zona pellucida. Gently wash the embryo through several drops of pre-equilibrated culture media before returning its actual culture drop for continued incubation (see Note 5).

22 3.1.3. Laser-Assisted Hatching of Blastocyst Stage Embryos (see Note 8)

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1. Prepare the assisted hatching dish with 10 ml droplets of micromanipulation medium. Quickly cover the drops with washed mineral or paraffin oil, making sure the drops are completely covered with oil. Warm the dish on a 37 °C surface for at least 15 min prior to micromanipulation (see Note 6). 2. Load the embryo(s) into the micromanipulation medium drops. 3. Focus on the embryo on high power (40×) and position the blastocyst (see Note 9). With the embryo in place, switch to the laser objective. 4. Focusing on the zona pellucida, fire the laser to create a 20 mM hole though the zona pellucida. 5. Using a pipet with an inner diameter of >250 mM, remove the embryo from the micromanipulation dish. Tighter pipets may squeeze the embryo and cause the blastomeres to escape the zona pellucida. Gently wash the embryo through several drops of pre-equilibrated culture media before returning its actual culture drop for continued incubation (see Note 5).

3.2. Embryo Culture

Embryo culture methods presented in previous chapters provide insight as to how embryologists can optimize conditions in their laboratory. Prolonged exposure to suboptimal conditions has been proposed to cause elevated rates of monozygotic twinning following extended embryo culture and blastocyst transfer (2, 4, 17, 24, 26). Despite this risk, the absence of tools to identify and select embryos with high reproductive potential at the cleavage stage, leads clinics to prefer blastocyst transfer, particularly for cases of single embryo transfer. If the theory proposed by Tummers et al. (31) that double implantation raises the risk of monozygotic twins is true, elective SET should be considered as method to reduce monozygotic twinning.

3.3. Elective Single Embryo Transfer

Programs must analyze their own data to determine which patients are good candidates for elective SET in their clinic. The methods below describe the criteria for SET at the University of Iowa Hospitals and Clinics. 1. Schedule all patients using oocytes from women under the age of 35 with six or more fertilized oocytes and women 35–37 with seven or more fertilized oocytes for a day 5 or blastocyst stage transfer. 2. Culture the embryos in groups of 3 or 4 in 50 ml drops of sequential culture media supplemented with 10 % protein under oil. All cultures are performed in 5–6 % CO2 in air at 37°C (see Note 10). 3. Approximately 114 h post-insemination, evaluate the embryos and assign quality grades. Embryos that have a blastocoele that

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comprises at least half the area of the blastocyst with the several cells in the inner cell mass, either tightly or loosely grouped, several cells forming a loose or cohesive epithelium in the trophectoderm are considered to be good quality. 4. Patients with at least one good quality blastocyst and no history of a previous failed fresh embryo transfer with the same oocyte source in the current clinic of care will undergo a single embryo transfer. If no good quality blastocysts are available, the patient may elect to transfer one or two embryos. 5. The embryo with the highest quality score is selected for transfer. Additional embryos that are graded as good quality expanded blastocysts may be cryopreserved per patient consent. Transfer of blastocysts that appear to have two inner cell masses or the inner cell mass appears to be “bridging” should be avoided. If there are no other good quality embryos available, the patient should be counseled regarding here elevated risk of monozygotic twins (see Notes 11 and 12).

4. Notes 1. As an extra precaution, draw a circle around the acidified medium drop on the underside of the dish to distinguish it from the manipulation medium drops. 2. Each manipulation medium drop is to be used once as the medium is acidified during assisted hatching (e.g., a second embryo should not be loaded into a drop that has already been exposed to the acidified medium). If assisted hatching is being performed without blastomere biopsy, more than one embryo may be loaded into the dish, but the duration of time the embryos are exposed to the acidified micromanipulation medium should be minimized. 3. The size of the hole is critical as holes 30 mM may allow blastomere loss in pre-compaction stage embryos (32). 4. The outer shell of the zona pellucida typically dissolves quickly, but the inner layer may prove to be elastic and resistant to the acidified medium. Expanding the hole in the inner layer may require mechanical disruption with the hatching pipet as exposure to excess acidified medium may compromise embryo development. 5. The embryologists handling embryos that have undergone assisted hatching must be mindful of the artificial gap in the

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zona pellucida throughout the remainder of the culture period and subsequent embryo transfer or cryopreservation. Always select a pipet with an inner diameter that provides liberal “clearance” for the embryo as it is being pipetted. 6. In contrast to using acidified medium, assisted hatching by laser does not change the chemical make-up and pH of the micromanipulation medium so the drops may be used multiple times for holding embryos. Furthermore, if embryo biopsy is not going to be performed, the embryologist may prefer to forego using a holding pipet. 7. Depending on the thickness of the zona pellucida, three to four pulses are typically required to cut completely through. 8. There have been no reports to date correlating an elevated risk of monozygotic twinning for blastocysts that have undergone assisted hatching. The accuracy and minimal trauma imposed by the laser make it the easiest method for assisted hatching the thin zona pellucida of the blastocyst stage embryo. 9. For trophectoderm biopsies, the blastocyst should be oriented with the ICM opposite the area to be hatched to avoid the risk that cells from the ICM are removed during biopsy. The ideal location for assisted hatching of blastocysts not undergoing biopsy is unknown. A recent study of assisted hatching in thawed blastocysts demonstrated that complete hatching was more likely if the zona was perforated near the inner cell mass (33). 10. Embryos are cultured in a cleavage medium that is glucose-, phosphate-free after the confirmation of fertilization (day 1). Unlike most sequential systems that move the embryos to a blastocyst culture medium designed for post-compaction stage embryos on day 3, we move day 3 embryos to a fresh drop of cleavage medium followed by transfer to blastocyst culture medium the morning of day 4. 11. Evidence of embryo splitting prior to embryo transfer on day 5 is rarely observed (34, 35). Meintjes et al. (35) reported establishment of a monozygotic triplet pregnancy after the transfer of a blastocyst with two inner cell masses. 12. The SET policy has been in place at our center for 6 years. During the first 3 years, the monozygotic twin pregnancy rate for patients undergoing SET was an alarming 5.8 % (9/155). During the most recent 3 years, we have been in a new laboratory and the monozygotic twin rate for SET has been 2.7 % (6/226). Only the space has changed. No other protocol or culture media changes were implemented between these two time periods. Our experience has been similar to that reported by Moeyeri et al. (36).

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References 1. Bulmer MG (1970) The biology of twinning in man. Oxford University Press, Oxford 2. Jain JK et al (2004) Monozygotic twins and triplets in association with blastocyst transfer. J Assist Reprod Genet 21:103–107 3. Blickstein I, Verhoeven HC, Keith LG (1999) Zygotic splitting following assisted reproduction. N Engl J Med 340:738–739 4. Milki AA et al (2003) Incidence of monozygotic twinning with blastocyst transfer compared to cleavage-stage transfer. Fertil Steril 79:503–506 5. Harkness UF, Crombleholme TM (2005) Twin-twin transfusion syndrome: where do we go from here? Semin Perinatol 29:296–304 6. Vela G et al (2011) Monozygotic pregnancies conceived by in vitro fertilization: understanding their prognosis. Fertil Steril 95:606–610 7. Dube J, Dodds L, Armson BA (2002) Does chorionicity or zygosity predict adverse perinatal outcomes in twins? Am J Obstet Gynecol 186:579–583 8. Schinzel AGL, Smith DW, Miller JR (1979) Monozygotic twinning and structural defects. J Pediatr 95:921–930 9. Hall J (2003) Twinning. Lancet 362:735–743 10. Carroll S et al (2005) Is zygosity or chorionicity the main determinant of fetal outcome in twin pregnancies. Am J Obstet Gynecol 193:757–761 11. Lee A et al (2010) The frequency and clinical significance of twin gestations according to zygosity and chorionicity. Twin Res Hum Genet 13:609–619 12. Sebire N et al (1997) The hidden mortality of monochorionic twin pregnancies. Br J Obstet Gynaecol 104:1203–1207 13. Paek B, Shields LE (2005) Twin-to-twin transfusion syndrome: diagnosis and treatment. Curr Women Health Rev 1:43–7 14. Mains L et al (2009) Sextuplets: an unusual complication of single embryo transfer. Fertil Steril 91(3):932.e1–932.e2 15. Van der Hoorn ML et al (2011) Dizygotic twin pregnancy after transfer of one embryo. Fertil Steril 95:805.e1–805.e3 16. Alikani M et al (1994) Monozygotic twinning in the human is associated with the zona pellucida architecture. Hum Reprod 9: 1318–1321 17. Vitthala S et al (2009) The risk of monozygotic twins after assisted reproductive technology: a systematic review and meta-analysis. Hum Reprod Update 15:45–55

18. Blickstein I (2005) Estimation of iatrogenic monozygotic twinning rate following assisted reproduction: Pitfalls and caveats. Am J Obstet Gynecol 192:365–368 19. Derom C et al (1987) Increased monozygotic twinning rate after ovulation induction. Lancet 1:1236–1238 20. Derom C et al (2006) High frequency of iatrogenic monozygotic twins with administration of clomiphene citrate and a change in chorionicity. Fertil Steril 85:755–757 21. Blickstein I, Keith LG (2007) On the possible cause of monozygotic twinning: lessons from the 9-banded armadillo and from assisted reproduction. Twin Res Hum Genet 10: 394–399 22. Slotnik R, Ortega J (1996) Monoamniotic twinning and zona manipulation: a survey of U.S. IVF centers correlating zona manipulation procedures and a high-risk twinning frequency. J Assist Reprod Genet 13:381–385 23. Yakin K, Balaban B, Urman B (2003) Risks of monochorionic pregnancies after assisted hatching. Fertil Steril 79:1044–1046 24. Skiadas C et al (2008) Risk factors associated with pregnancies containing a monochorionic pair following assisted reproductive technologies. Hum Reprod 23:1366–1371 25. Menezo Y, Sakkas D (2002) Monozygotic twinning: is it related to apoptosis in the embryo? Hum Reprod 17:247–248 26. Behr B et al (2000) Blastocyst-ET and monozygotic twinning. J Assist Reprod Genet 17:349–351 27. Chang HJ et al (2009) Impact of blastocyst transfer on offspring sex ratio and the monozygotic twinning rate: a systematic review and meta-analysis. Fertil Steril 91:2381–2390 28. Kawachiya S et al (2011) Blastocyst culture is associated with an elevated incidence of monozygotic twinning after single embryo transfer. Fertil Steril 95(6):2140–2142. doi:10.1016/j.fertnstert.2010,12.018 29. Sills ES, Tucker MJ, Palermo GD (2000) Assisted reproductive technologies and monozygous twins: implications for future study and clinical practice. Twin Res 3: 217–223 30. Knopman J et al (2009) Monozygotic twinning: an eight-year experience at a large center. Fertil Steril 94:502–510 31. Tummers P, De Sutter P, Dhont M (2003) Risk of spontaneous abortion in singleton and twin pregnancies after IVF/ICS. Hum Reprod 18:1720–1723

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Chapter 23 Embryo Culture and Epigenetics Brenna A. Market Velker, Michelle M. Denomme, and Mellissa R.W. Mann Abstract During preimplantation development, major epigenetic reprogramming occurs, erasing gametic modifications, and establishing embryonic epigenetic modifications. Given the plasticity of these modifications, they are susceptible to disruption by assisted reproductive technologies, including embryo culture. The current state of evidence is presented for the effects of embryo culture on global DNA methylation and histone modifications, retroviral silencing, X-inactivation, and genomic imprinting. Several salient points emerge from the literature; that culture in the absence of other procedures can lead to epigenetic perturbations; that all media are suboptimal; and that embryo response to in vitro culture is stochastic. We propose that embryos adapt to the suboptimal environment generated by embryo culture, including epigenetic adaptations, and that “quiet” embryos may be the least epigenetically compromised by in vitro culture. Key words: Embryo culture, Epigenetics, Endogenous retroviral silencing, X inactivation, Genomic imprinting, DNA methylation, Histone modifications

1. Introduction The field of epigenetics encompasses the study of heritable and reversible modifications of chromatin that influence the accessibility of genes and regulate gene transcription (1). Alterations in gene expression are not brought about by changes to the DNA sequence, yet the epigenetic information affecting gene activation or repression ultimately allows for variation in cellular phenotypes. Epigenetic mechanisms include DNA methylation and histone posttranslational modifications. These mechanisms play important roles in the establishment of cellular memory and differentiation during embryogenesis (2), and given their plasticity, are susceptible to disruption by assisted reproductive technologies (ARTs).

Gary D. Smith et al. (eds.), Embryo Culture: Methods and Protocols, Methods in Molecular Biology, vol. 912, DOI 10.1007/978-1-61779-971-6_23, © Springer Science+Business Media, LLC 2012

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Fig. 1. (a) Epigenetic reprogramming during mammalian preimplantation development. Following fertilization, the maternal and paternal genomes must be epigenetically reprogrammed from a gametic to an embryonic state. Reprogramming is initiated in pronuclear stages and continues throughout preimplantation development. Maternal gametic genome (red ), paternal gametic genome (blue), and embryonic genome (green). (b) Histone modifications. Protamines are actively removed from the paternal genome (shades of blue) immediately following fertilization. Activating histone modifications are set in the early pronuclear stages (H4Ac/H3K4me1), which are followed by acquisition of repressive histone modifications (H3K4me3/H3K9me2/HeK27me2/3). Both activating (H4Ac/H3K4me1) and repressive (H3K9me2/3/H3K27me1/H4K20me3) histone modifications are abundant throughout the maternal genome (pink) during pronuclear stages. (c) DNA Methylation. The paternal genome (blue) undergoes active global demethylation immediately following fertilization, while the maternal genome (red ) is passively demethylated through subsequent cleavage divisions. Line, ERV-L, and etn (orange) follow a similar demethylation process, while IAPs (yellow ) and imprinted loci (green) are protected from demethylation events. (d) X-inactivation. A cascade of repressive modifications occurs on the paternal X chromosome in female preimplantation embryos, beginning with paternal-specific Xist expression.

During mammalian development, there are two major phases of epigenetic reprogramming (2, 3). During gametogenesis, reprogramming consists of erasure of previous epigenetic marks that restore totipotency, followed by the establishment of sex-specific epigenetic marks. Following fertilization, major epigenetic reprogramming occurs for a second time, and includes alterations in histone posttranslational modifications and DNA methylation (Fig. 1). 1.1. Histone Modifications

The nucleosome is the basic chromatin unit, containing doublestranded DNA wrapped around an octamer of core histone proteins (4). Chromatin is a dynamic structure that undergoes

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alterations in its packaging, affecting the accessibility of underlying genes (5). Heterochromatin is a highly condensed structure that results in gene silencing, while euchromatin is a less compacted configuration supporting gene expression. Switching between active and repressive chromatin states is facilitated by covalent posttranslational modifications of histone tails (5). Histone tail modifications, such as acetylation and methylation, confer either activating or repressive functions, depending on the type of modification and the amino acid residue that is modified. Shortly after fertilization, the paternal genome undergoes extensive remodeling that includes an exchange of protamines for histones, and acquisition of active histone modifications, including histone H4 acetylation (H4Ac) and histone H3 lysine 4 methylation (H3K4me1) (6) (Fig. 1). In early embryos, the paternal genome also acquires repressive histone modifications, including histone 3 lysine 9 and lysine 27 methylation (H3K9me2, H3K27me2, and H3K27me3). By comparison, the maternal genome possesses both active (H4Ac, H3K4me1) and repressive histone modifications (H3K9me2, H3K9me3, and H4K20me3). These covalent modifications are proposed to initiate the transcriptionally repressed state that coincides with embryonic genomic activation. This potential for chromatin bivalency, where both activating and repressive marks occupy the same stretch of chromatin, is likely a major factor in establishing the correct gene expression profile for embryonic development (7). Switching between active and repressed chromatin states is also brought about by incorporation of specialized histone variants and chromatin remodeling proteins. One histone variant present during early mouse development is histone 3.3 (H3.3), which is first detected in the paternal pronucleus after the protamine to nucleosome exchange, and then at the 2-cell, 4-cell, and blastocyst stages in correlation with waves of embryonic gene activation (8). H3.3 is exchanged for H3 to promote transcriptionally active regions (9). In addition, chromatin remodeling proteins can change chromatin structure in a non-covalent manner. Functioning antagonistically, Polycomb group proteins act to establish a silenced chromatin state via polycomb repressive complexes (PRC1 and PRC2), while Trithorax group proteins act to promote a permissive chromatin state (10). In the mouse zygote, PRC1 and PRC2 complexes are localized to the maternal pronucleus at earlier pronuclear stages (PN0–3), and are observed at the paternal pronucleus by the latest pronuclear stage (PN5), initiating the repressive pathways for maternal and paternal constitutive heterochromatin in the embryo (11). Thus, extensive epigenetic reprogramming occurs in histone and chromatin remodeling during early development.

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1.2. DNA Methylation

DNA methylation is the covalent modification of cytosines within CpG dinucleotides that is typically associated with gene silencing. DNA methyltransferases, DNMT3A, DNMT3B along with DNMT3L, act as de novo methyltransferases, establishing methylation marks. Acquisition of DNA methylation occurs in the developing gametes and is acquired differentially between the two parental genomes (12, 13). By comparison, DNMT1 is responsible for maintaining DNA methylation throughout preimplantation development through its recognition of hemi-methylated DNA. An oocyte-specific isoform, DNMT1o, is present in the mature oocyte, and localizes to the maternal pronucleus post-fertilization (14). While DNMT1o is present during all stages of preimplantation development, it is predominantly cytoplasmic, except at the 8-cell stage when it is translocated to the nucleus (15, 16). Therefore, its function in maintaining DNA methylation during preimplantation development is not completely clear. A somatic isoform, DNMT1s, which localizes to the nucleus throughout preimplantation development, is responsible for DNA methylation maintenance at repetitive elements and imprinted genes (17). Paradoxically, while the fertilized egg is highly enriched for DNMT proteins, there is a wave of demethylation during preimplantation development that erases gamete-specific methylation patterns and ensures the totipotency of the early embryo (3). The paternal genome is actively demethylated within hours after fertilization, while the maternal genome is passively demethylated during early cleavage divisions in a replication-dependent manner (Fig. 1). The end result is that the two parental genomes undergo extensive changes in global methylation during preimplantation development. Postimplantation, de novo methylation gradually increases in accordance with cellular differentiation. In addition to these generalized epigenetic gene regulatory mechanisms, there are three specialized epigenetic phenomena undergoing dynamic regulation during early embryonic development. These phenomena are endogenous retroviral silencing, X chromosome inactivation, and genomic imprinting.

1.3. Endogenous Retroviral Silencing

Mammalian genomes are highly enriched for repetitive sequences, of which the vast majority are endogenous retroviral elements (18). To reduce the deleterious effects of retrotransposons, the host epigenetically silences these elements. As such, preimplantation development is a critical period with dynamic changes in expression and DNA methylation of retroviral elements. Mouse endogenous retroviral-like element (ERV-L) and interstitial A particles (IAP) are expressed from the 2-cell stage and are repressed by the blastocyst stage (19, 20). With regard to DNA methylation, some retro-elements are demethylated similar to the majority of the genome including human L1, mouse ERV-L, and mouse early retrotransposons (etn), while IAPs remain heavily methylated and

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protected from this demethylation event (14, 21, 22) (Fig. 1). DNMT1 is the methyltransferase responsible for maintenance methylation at IAPs (14, 23). Given the dynamic nature of epigenetic retroviral regulation, a mouse model system, agouti viable yellow (Avy), has been used to study environmental exposure. Avy mice differ from wildtype mice, as they possess a transposable IAP element upstream of the agouti gene, which contains an alternate, constitutive promoter (24, 25). Hypomethylation of the retrotransposon allows for ectopic agouti transcription, which leads to yellow fur as well as adult-onset obesity, diabetes, and tumor development (26, 27). Hypermethylation of the transposable element silences ectopic expression, leading to wildtype agouti coat color and protection against associated pathology. These mice are ideal for environmental studies as genetically identical Avy mice can exhibit an array of coat color phenotypes based on insult-induced epigenetic alterations during early development. For example, maternal exposure to deleterious environmental toxicants, such as bisphenol A, can reduce methylation levels within the fetal genome, causing hypomethylation at the Avy gene (28). Conversely, maternal nutritional supplementation with methyl donors (folic acid, betaine, vitamin B12, and choline) can protect against loss of methylation, permitting hypermethylation of Avy gene (29). A second locus of interest is the Axin1-Fused (Axin1Fu) gene. In this case, a transposable IAP element, which possesses a cryptic promoter, is located with intron 6 of the Axin1 gene (30). Hypomethylation of the Axin1Fu allele permits mutant transcripts to be expressed, generating the Axin1Fu kinked tail phenotype, while hypermethylation of the Axin1Fu allele silences expression from the cryptic promoter, preventing tail kinks (31). These mice are also well suited for environmental studies as genetically identical Axin1Fu mice can exhibit an array of kinked tail phenotypes based on environmental insult during early development (31, 32). Thus, the Avy and Axin1Fu mouse models represent excellent systems to determine the epigenetic effects of environmental insult during embryo development, including embryo culture. 1.4. X-Inactivation

Dosage compensation in mammals is achieved by X chromosome inactivation, whereby XX females silence one sex chromosome to enable equal X-linked gene expression to that of XY males. The noncoding X-inactive specific transcript (Xist) RNA located within the X chromosome inactivation center is responsible for this process (33). During preimplantation development in the mouse, Xist is transcribed from the paternal X and coats the inactive X chromosome, initiating a cascade of epigenetic modifications that result in silencing of the entire chromosome (34). The propagation of X inactivation along the chromosome involves a hierarchy of epigenetic changes that are initiated with Xist expression (35–37)

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(Fig. 1). Around the 8-cell stage, active histone modifications (H3K9Ac, H3K4me2) are lost. Chromatin remodeling complexes are then recruited to the paternal X chromosomes around the 16-cell stage, conferring the repressive histone modification histone 3 lysine 27 trimethylation (H3K27me3) (35–37). At this time, the repressive histone variant macroH2A is also incorporated into the paternal X chromosome. At the 32-cell stage, the repressive histone modification H3K9me3 is acquired on the paternal X (35–37). At the blastocyst stage, the trophectodermal lineage retains this imprinted form of X chromosome inactivation, while in the inner cell mass, the paternal X is reactivated, and random X chromosome inactivation is initiated (36, 38). 1.5. Genomic Imprinting

Genomic imprinting is an epigenetic phenomenon whereby certain genes are expressed exclusively from one parental allele (33). These imprinted genes often reside within domains, where multiple imprinted genes are coordinately regulated in cis by DNA elements known as imprinting centers. Genomic imprinting is the result of both acquisition of imprinting marks during gametogenesis, whereby oocytes acquire maternal-specific DNA methylation imprints and sperm obtain paternal-specific DNA methylation imprints, and of maintenance of these marks throughout development. During preimplantation development, a wave of demethylation erases gamete-specific methylation from the maternal and paternal genomes. However, DNA methylation imprinting marks are excluded from this demethylation event; the methylated allele remains methylated and protected from demethylation, while the unmethylated allele remains unmethylated (3) (Fig. 1). Maintenance of these methylation marks through preimplantation development requires expression of DNMT1 (14, 17, 39, 40), as well as changes in histone modifications, including H3K9me2 and H3K27me3 enrichment on the repressed allele (41). Of significance is that many imprinted genes play critical roles in the development of the embryo, and their misregulation has been linked to the development of human imprinting disorders, such as Angelman syndrome (AS) and Beckwith–Wiedemann syndrome (BWS). AS is an imprinting disorder characterized by a severe mental retardation, seizures, little or no speech, an abnormal gait and frequent smiling/laughing behavior (42), while BWS is an overgrowth disorder characterized by macroglossia, abdominal wall defects, birth or postnatal weight above the 90th percentile, neonatal hypoglycemia, and an increased incidence of Wilm’s tumor (43). Given the dynamic regulation of endogenous retroviral silencing, X chromosome inactivation, and genomic imprinting during early embryonic development, these specialized epigenetic mechanisms have been the focus of studies involving assisted reproduction, with genomic imprinting the most widely investigated.

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1.6. Embryo Culture

Given that epigenetic mechanisms mediate interactions between the stable intrinsic genome and variable extrinsic environment, many studies have investigated the effects of environmental insult on epigenetic gene regulation. As such, scientists and clinicians alike have questioned the environmental impact of embryo culture on epigenetic programming. While we focus on epigenetic effects of embryo culture, the following caveat should be noted. In many papers, ARTs and their observed effects are often grouped together under the umbrella of “potential consequences of ARTs”, making it nearly impossible to separate effects of embryo culture from those of other techniques. Thus, one must bear in mind that the described effects of embryo culture may be attributed to more than one procedure. Given the numerous and varied forms of ARTs currently employed in research and human clinics, investigation of distinct procedures should be a clear objective of the field moving forward.

1.6.1. Epigenetic Effects of Embryo Culture

It has been proposed that embryo culture may have a global effect on DNA methylation patterns, chromatin remodeling, and gene expression. To assess the effects of in vitro culture on DNA demethylation, the dynamics of genome demethylation were assayed using immunocytochemistry with 5-methylcytosine antibodies (44, 45). Comparison between in vitro cultured and in vivo-derived mouse and rat embryos revealed a decreased ability of cultured embryos to efficiently demethylate the paternal genome, indicating that embryo culture may interfere with the earliest epigenetic reprogramming of the gamete-derived genomes. Global changes in DNA methylation have also been analyzed. Recently, the effect of assisted reproduction on DNA methylation was assessed at more than 700 genes in placenta and cord blood from children of assisted and unassisted conceptions using a sitespecific CpG methylation assay (46). A significant change in DNA methylation was observed with lower mean methylation levels in the placenta, and higher mean methylation levels in cord blood from children conceived in vitro when compared to children conceived naturally, indicating the broad effects of ART on DNA methylation. To date, only one study has examined global changes in histone modifications. Using immunohistochemistry, H4 acetylation, H3K9 methylation, or H3S10 phosphorylation was examined in mouse embryos from the 1-cell to blastocyst stage following in vitro fertilization (47). No difference was found between in vitro cultured and in vivo-derived mouse embryos for these modifications. Given the limited nature of immunofluorescence analyses in detecting specific changes that may occur in a globally modified background, future studies should be directed towards high throughput sequence-based assays. However, the limited material available in individual preimplantation embryos makes studies utilizing these current techniques difficult at best.

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In addition to global epigenetic changes, effects of embryo culture on endogenous retroviral silencing, X chromosome inactivation, and genomic imprinting have also been investigated. Two endogenous retroviral mouse models, Avy and Axin1Fu, have been employed to study the environmental impact of embryo culture on retroviral silencing. In these studies, effects of embryo culture during preimplantation development were queried by examining the phenotypic effects of postnatal Avy and Axin1Fu expression. In vitro culture was found to increase the agouti yellow and tail kink phenotypes, respectively (48, 49). This indicates that embryo culture interferes with maintenance of epigenetic silencing of IAPs, allowing an active state of the IAP allele to be propagated. In the case of the agouti yellow phenotype, the Avy allele displayed a corresponding loss of DNA methylation, while for the kinked tail phenotype, the Axin1Fu allele exhibited H3K4 dimethylation and H3K9 acetylation, two hallmarks of active chromatin (48, 49). These results again demonstrate the adverse impact of embryo culture on epigenetic gene regulation. With respect to X chromosome inactivation and embryo culture, very few studies have been performed. In bovine, increased XIST transcript abundance was observed in in vitro-cultured compared to in vivo-derived embryos (50, 51). In the mouse, Xist was inappropriately expressed from the sole, maternally derived X chromosome in male midgestation placenta following preimplantation in vitro culture, perhaps indicating that the imprinted form of X-inactivation is disrupted by embryo culture (52). In contrast to X chromosome inactivation, a significant body of evidence has emerged regarding the aberrant regulating of genomic imprinting following in vitro culture of mouse, human, and livestock embryos. The first study to demonstrate a link between embryo culture and genomic imprinting reported biallelic expression of the maternally expressed H19 gene in a subset of pooled, cultured mouse embryos (53). A similar study examining H19 imprinted expression and methylation revealed aberrant biallelic H19 expression and loss of paternal-specific methylation in cultured mouse blastocysts (54). Since then, it has been determined that in vitro preimplantation culture of mouse embryos results in biallelic expression of the H19 gene and loss of imprinted methylation at the H19, Snrpn, and Peg3 genes in blastocyst stage embryo (52–58). Subsequent observations of postimplantation embryos indicate that epigenetic alterations induced by embryo culture persist. At day 9.5, following embryo culture and embryo transfer, loss of methylation and biallelic expression was observed for H19, Snrpn, Peg3, and Kcnq1ot1 in extraembryonic tissues, indicating that imprinting perturbations are inherited through to midgestation, long after embryos have been removed from in vitro culture (52, 53, 59).

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Genomic imprinting may also be affected by in vitro manipulation of early embryos during ARTs for the treatment of infertility/ subfertility in humans. Increased incidence of the human imprinting disorders AS and BWS has been reported with the use of ARTs (60–68). More specifically, the incidence of Angelman syndrome in the general population is approximately one case per 16,000 births, with only 5% of these cases related to imprinting abnormalities (42, 60). As the prevalence of AS is low, large-scale studies containing sufficient numbers of patients have been difficult to achieve. However, seven cases of AS following the use of ARTs have been reported to date, five of which displayed imprinting abnormalities (71%) (60, 65, 68, 69). This is a significantly higher proportion in the ART population compared with the non-ART population. The specific epigenetic defect of AS has been mapped to the Snrpn imprinting center. Loss of methylation at the paternal Snrpn imprinting center results in overexpression of the paternally expressed genes of this domain and loss of expression of the maternally expressed genes. BWS is a second imprinting disorder associated with ARTs. BWS affects about one in 13,700 children (43). As with AS, parents of children with BWS are more likely to have undergone fertility treatments than the general population, and a higher incidence of BWS is seen in ART children compared with naturally conceived children (61–63, 65, 67, 69). Two imprinted defects have been linked to BWS. With respect to the first, the maternal H19 imprinted domain acquires a paternal epigenotype. Gain of H19 imprinting center methylation results in silencing of H19 expression and activation of Igf2 expression. The second imprinting defect occurs at the Kcnq1ot1 domain and results from loss of methylation on the maternal Kcnq1ot1 imprinting center with ensuing biallelic expression of the long noncoding RNA Kcnq1ot1, and silencing of Kcnq1 and Cdkn1c. The prevalence of ARTs is approximately 4–9 times greater in BWS children compared to the number of ART children in the general population. Taken together, ARTs may impose inherent risk for normal imprinted gene regulation. Bovine and ovine animals produced through in vitro fertilization and other forms of ARTs display an increased frequency of “Large Offspring Syndrome”, which is characterized by a variety of abnormalities including increased birth weight, organomegaly, polyhydraminos, hydrops fetalis, immunological defects, increased fetal/ neonatal death as well as numerous skeletal and placental defects (70, 71). Interestingly, the phenotypes observed with Large Offspring Syndrome are similar to those observed in Beckwith– Wiedemann syndrome. Importantly, perturbation of the same loci involved in BWS is observed in calves with Large Offspring Syndrome. KCNQ1OT1 is hypomethylated with a corresponding increase in KCNQ1OT1 expression and decrease in CDKN1C

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expression (72). Large offspring sheep also exhibit loss of imprinting for the growth inhibiting Insulin-like growth factor 2 receptor gene (73). Furthermore, loss of maternal-specific SNRPN methylation and activation of normal silent maternal SNRPN allele was found in placenta from in vitro fertilized and cultured bovine embryos, similar to AS in humans (74). The similarities in ART-induced epigenetic errors in humans and in animal models, where subfertility is not a confounding issue, reinforces the notion that manipulation of the early embryo can lead to epigenetic perturbations which result in long-term consequences for the offspring. One interesting observation from the above studies is that not all embryos, and not all imprinted loci in these embryos, are affected by in vitro culture. This indicates that the response of embryos to in vitro culture is stochastic. Analyses at the individual embryo level have confirmed this (52, 57, 58). Similar observations have been recently reported in ART-conceived children with BWS (66, 75). Imprinting defects at multiple imprinted loci in addition to the Kcnq1ot1 gene were present in BWS patients whose parents had undergone some form of ART. These data suggest that developmental defects or abnormal growth in ART children might be caused by variable combinations of epigenetic perturbations at imprinted genes. The mechanism by which some embryos are able to compensate for the suboptimal preimplantation environment while others are not is unknown. However, it has been hypothesized that modulation of the expression of a group of imprinted genes (known as the “imprinted gene network”) may play a role (57). All together, these studies have confirmed maintaining preimplantation embryos outside the female reproductive tract using embryo culture affects genomic imprinting at multiple loci. In humans, infertility/subfertility may also be a contributing factor in aberrant imprinted gene regulation. Thus, it is clear that embryo culture constitutes an environmental insult to epigenetic gene regulation. However, the specific aspect of embryo culture most responsible for this epigenetic misregulation has yet to be determined. 1.6.2. Epigenetic Effects of Culture Media

Many studies have attempted to show superiority of one media or another, with respect to various measures of developmental competence (76–78). With respect to gene expression, a comparison was made between embryos cultured in KSOM augmented with amino acids (KSOMaa) with those cultured in Whitten’s medium. Global expression analysis revealed that a larger number of genes were misexpressed after culture in Whitten’s medium compared with culture in KSOMaa (79), suggesting aberrant epigenetic regulation of gene expression in Whitten’s medium. With regard to X chromosome inactivation, Xist expression has been analyzed in midgestation placental tissues from male embryos subjected to preimplantation in vitro culture. Culture in

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Whitten’s medium produced a greater number of embryos with ectopic Xist expression from the sole active X chromosome compared to culture in KSOMaa (52). The effects of various culture media on genomic imprinting have also been evaluated. In the mouse model, while KSOMaa was better able to maintain genomic imprinting than Whitten’s medium, KSOMaa has also been shown to cause disruptions in H19, Snrpn, Peg3, and Ascl2 imprinting at the blastocyst stage, and H19, Snrpn, Igf2, Kcnq1ot1, Cdkn1c, Mkrn3, Ascl2, Zim1, and Peg3 imprinting at postimplantation stage (day 9.5) (52, 54, 59). In addition, at the blastocyst stage, M16 medium was shown to cause greater perturbation of H19 imprinting than G1.2/G2.2 (57). Human tubal fluid medium (HTF) caused loss of H19 imprinting including aberrant histone modifications with an increase in H3K4 dimethylation on the paternal H19 allele and an increase in H3K9 trimethylation on the maternal H19 allele (56). In bovine, alterations in imprinting of Snrpn have been associated with the nonsequential SOF medium (74) as well as Sgce and Ata3 in the sequential vitro cleave/vitro blast medium (80). In humans, a recent study reported loss of methylation at the H19 locus in ~19% of a cohort of human embryos cultured in cleavage medium (81). As imprinting perturbations arise from multiple media systems, it suggests that all media are suboptimal in supporting normal genomic imprinting. Of note however, is that all the above studies employed ovarian stimulation to retrieve embryos prior to culture. To assess the effects of various culture media on genomic imprinting in the absence of other ARTs, we performed a comparison of six embryo culture systems, Whitten’s; KSOMaa; HTF; Global; Preimplantation 1/Multiblast (P1/MB); and G1v5PLUS/ G2v5PLUS (G1.5/G2.5), in the mouse model system. Blastocysts cultured in each of the six culture systems experienced a loss of H19, Snrpn, and Peg3 imprinting (58). With regard to imprinted methylation, better preservation of H19, Snrpn, and Peg3 imprinted methylation was observed in embryos cultured in KSOMaa compared to the other media systems where a greater loss of imprinted methylation was observed. In addition, a genedependent response was observed with certain media supporting higher levels of imprinted methylation at some but not at all loci. More specifically, embryos cultured in KSOMaa, Global and P1/ MB displayed higher levels of H19 imprinted methylation than Whitten’s, HTF and G1.5/G2.5. For Snrpn, KSOMaa, and Global better maintained imprinted methylation than the other media systems, and for Peg3, levels of imprinted methylation for KSOMaacultured embryos were best, while Whitten’s and G1.5/G2.5 cultured embryos showed the greatest loss of imprinted methylation. Of note is that sequential media systems did not seem to confer an advantage with respect to maintenance of genomic

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imprinting compared to their single step counterparts, nor did medium renewal. Our results confirmed the observation that in vitro culture of preimplantation embryos alone leads to alterations in imprinted gene regulation. 1.6.3. Epigenetic Effects of Serum

The addition of serum to embryo culture media, although frequently used for somatic cell culture, has been shown to be detrimental to the development of the early embryo resulting in decreased developmental potential (82, 83). Two studies in mouse have addressed the effects of serum on genomic imprinting. The addition of serum to the culture medium was shown to have severe consequences with respect to the epigenetic integrity of the early embryo, resulting in alterations in the expression and methylation of various imprinted genes (H19, Igf2, Grb7, Grb10, and Peg1/Mest), a faster rate of development to the blastocyst stage, and long-term behavioral consequences (55, 84). In addition, the use of serum is highly associated with the development of Large Offspring Syndrome (71, 85, 86). As serum contains undefined growth factors and other macromolecules, the mechanism of these perturbations remains unknown. Thus, the use of serum in embryo culture is discouraged.

1.6.4. Epigenetic Effects of Oxygen Tension

In the female reproductive tract, embryos are maintained in a relatively hypoxic environment, with oxygen levels around 5% (87, 88). Culture of mammalian embryos in low oxygen improves many aspects of embryo development (89–94). These results indicate that the overall epigenetic landscape of embryos cultured in 5% oxygen is more similar to in vivo-derived embryos than those cultured in 20% oxygen (95). The effect of various oxygen concentrations on genomic imprinting in particular has not been well studied. Overall, expression levels of a subset of imprinted genes were not noted to be different in embryos cultured in high levels of oxygen post-compaction compared to those cultured in low levels. However, this investigation was not able to distinguish whether expression of these genes was from the maternal allele, paternal allele, or both (96). With the clear advantage of low oxygen tension with respect to overall embryo development, in vitro culture in low oxygen is highly recommended.

1.6.5. Multiple ARTs

As noted above, many studies examining the epigenetic effects of embryo culture have employed multiple ARTs. With this in mind, several groups have conducted research to distinguish between the effects of various ARTs. In mouse, one study tested the effects of superovulation alone, superovulation and in vitro culture following natural fertilization, and superovulation, in vitro fertilization, and in vitro culture (57). The group subjected to all three procedures showed the greatest disruption to H19 imprinting in individual blastocysts. Similarly, we also observed that increased loss of

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H19 imprinted expression as a result of embryo culture that was exacerbated by the use of superovulation (58). Finally, a study by Rivera et al. demonstrated an increase in the number of embryos with biallelic H19, Cdkn1c, Kcnq1, Snrpn, Kcnq1ot1, Peg3, and Mkrn3 expression following superovulation with embryo transfer compared to controls, with a further increase in biallelic expression following embryo culture combined with superovulation and embryo transfer (59). Together, these studies demonstrate an additive effect with combined ART procedures producing a greater chance of epigenetic perturbations than single interventions. In a recent study, we examined the effects solely of superovulation on genomic imprinting in a mouse model. As superovulation is thought to primarily affect oocyte development, effects were expected to be limited to maternal alleles. In addition to aberrant maternal-specific methylation, we found that paternal H19 methylation was perturbed by superovulation (97). This suggests that superovulation affects maternal-effect gene products subsequently required for imprint maintenance during preimplantation development. Our new working model is that the effects of superovulation and embryo culture converge in the preimplantation embryo on the same epigenetic pathway that is supported by oocyte-specific product(s). In fact, it is not unreasonable that a number of environmental insults, such as oxygen tension, serum, waste byproducts, and embryo transfer may perturb the same epigenetic pathway in a stochastic manner. This would be consistent with ART-associated BWS children where variable ART procedures were reported to be used, including embryo culture media (varied in type, and in glucose, amino acid and human sera albumin content), day of transfer, and specific ART method used (in vitro fertilization, intracytoplasmic sperm injection), in addition to the cause of infertility (62, 69). This is consistent with a recent study where no differences were observed in global DNA methylation and chromatin organization in human embryos generated via in vitro fertilization or intracytoplasmic sperm injection (98). Furthermore, a recent study of H19 imprinted methylation examined intra- and interindividual variation in DNA methylation in in vitro or in vivo conceived children (99). Substantial variation was found in allelespecific H19 methylation, with greater aberrant methylation and greater variability in abnormal methylation in the in vitro group. Furthermore, consistent with animal model studies, placenta harbored greater perturbation than cord blood or tissue. We interpret these results to mean that there is stochastic imprinting perturbation at the individual cell level following in vitro conception. If embryo response to in vitro culture is stochastic, then the leading question is which embryo will be most compromised by suboptimal culture. In our most recent study, we investigated rates of development and loss of genomic imprinting (100). We reasoned that as culture embryos exhibit delayed development, that

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slower developing embryos would exhibit greater perturbation in imprinted gene regulation than their counterparts that possess faster rates of development. Contrary to our expectations, embryos with faster rates of development experienced the greatest loss of imprinting (100). Furthermore, expression of two markers of metabolic activity revealed that the fastest developing embryos were most divergent from in vivo-derived controls. Our results are consistent with the Quiet Embryo hypothesis which states that those embryos which are able to maintain a relatively quiet or low level of metabolism are most viable (101, 102). Therefore, embryos that adapt the least to their suboptimal environment, including through epigenetic alterations, will likely develop in the most normal fashion. Thus, development of culture media, which minimize the need for increased metabolic activity, will likely have a corresponding decrease in epigenetic perturbations and thus, increased overall health of the embryo.

2. Conclusions In conclusion, all of the above studies on epigenetic gene regulation in early embryo development reinforce the point that preimplantation development is a dynamic period of epigenetic programming and suboptimal embryo culture produces epigenetic adaptations. While it is possible to generate blastocysts that appear morphologically normal, some in reality are epigenetically compromised. It is up to those pursuing research in these fields to first determine accurate, repeatable, and noninvasive techniques to detect these compromised embryos, as well as to continue to improve current techniques to minimize the amount of adaptation that embryos will require to survive and grow outside the female reproductive tract.

3. Materials The following are solutions for the combined imprinted expression and methylation analysis for a single blastocyst stage embryo. Prepare all solutions with double-distilled autoclaved water. Use filter tips and nonstick surface microfuge tubes. Wash all pipetmen and equipment with RNAse Away (Invitrogen, Burlington, Canada). To reduce the chance of DNA and RNA contamination, change gloves often. Wipe all pipetmen, tubes, tip boxes and other reagents with RNAse Away after each change of gloves. Keep tubes angled away when open, and recap all tubes when not in use. 1. Dynabead mRNA Direct Micro kit (Dynal Biotech, Invitrogen, Burlington, Canada).

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2. Ready-To-Go PCR beads (GE, Healthcare, Quebec, Canada). 3. 10% Igepal: 1 ml of Igepal (Sigma, Oakville, Canada) in 9 ml H2O. 4. 3 M NaOH: 2.4 g NaOH in 20 ml H2O. 5. 0.1 M NaOH: 0.5 ml 3 M NaOH in 14.5 ml H2O. 6. 0.3 M NaOH: 1 ml of 3 M NaOH in 9 ml H2O. 7. ITT Buffer: 50 μl of 10% Igepal and 50 μl of 10% Tween 20 in 50 ml of TE (pH 7.5). 8. 2% Low melting point (LMP) agarose: 20 mg Agarose in 1 ml H2O. Dissolve at 70°C, then incubate at 90°C until use. 9. 2.5 M Bisulfite solution Part 1 (see Note 1): 3.8 g Sodium bisulfite, 5.5 ml H2O, 1 ml 3 M NaOH. Dissolve at 50°C. 10. Bisulfite solution Part 2: 110 mg Hydroquinone, 1 ml H2O. Dissolve at 50°C. When fully dissolved, mix bisulfite solution Parts 1 and 2. Store away from light at room temperature until use.

4. Methods 4.1. Prewash Dynabeads

1. Vortex Dynabeads on medium speed to resuspend. Change gloves. 2. Label one 0.2 ml thin-walled PCR tube per sample plus an additional tube for a negative control. 3. Transfer 10 μl of Dynabeads to each 0.2 ml thin-walled PCR tube. 4. Place into magnetic particle concentrator (MPC). Remove supernatant. 5. Add 100 μl Dynabead lysis buffer to each sample. 6. Vortex on low for 5–10 s, place on MPC, then remove buffer. Repeat.

4.2. Embryo Lysis

1. Retrieve individually frozen embryos from storage at −80°C (see Note 2). 2. Centrifuge briefly in microfuge to ensure embryo is at bottom of tube (~5 s at 9503–16060´g). 3. Add 10 μl of Dynabead lysis buffer to each sample. 4. Prior to adding embryo, place in MPC, and remove lysis buffer from prewashed Dynabeads from step 6 above. 5. Transfer entire contents of lysed embryo sample from step 3 to prewashed Dynabeads.

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6. Mix gently by flicking, then centrifuge briefly at 1,520´g in microfuge. 7. Incubate with slow agitation on vortex for 5–10 min at room temperature to allow for hybridization of mRNA to Dynabead. 8. Centrifuge briefly at 1,520´g. 9. Place mRNA-Dynabead tube in MPC. Remove supernatant containing DNA to original embryo tubes, taking care not to remove any mRNA-Dynabeads. 10. Add 200 μl of Dynabead Wash Buffer A to mRNA-Dynabeads. Place in MPC and set aside until 3.3 RNA isolation. 11. Centrifuge tubes containing DNA/supernatant for 5–10 s in microfuge at 16,060´g to remove bubbles. 12. Add 1 μl Protease K (Sigma, Oakville, Canada) and 1 μl of 10% Igepal (see Note 3) to each DNA/supernatant tube. 13. Centrifuge briefly in microfuge at 16,060´g to remove any bubbles. 14. Add 300 μl of DNAse-, RNAse-, and protease-free mineral oil to each tube (see Note 4). 15. Lyse embryo by incubating for 1 h in waterbath at 50°C. 4.3. RNA Isolation

1. Retrieve mRNA-Dynabead tube containing 200 μl of Wash Buffer A. 2. Vortex on low speed for approximately 5 s. Centrifuge briefly. Place in MPC, and then remove Wash. 3. Repeat washing step with Wash Buffer A once, and Wash Buffer B three times.

4.4. Reverse Transcription and Generation of a Solid-Phase cDNA Library (see Note 5)

1. Prepare reverse transcription by mixing 2 μl 5× First Strand Buffer, 1 μl 0.1 M DTT, 0.5 μl 10 mM dNTP, 0.5 μl 40 U/μl RNaseOut (Invitrogen, Burlington, Canada), 0.25 μl Superscript II (Invitrogen, Burlington, Canada), 5.75 μl H2O, for a 10 μl reaction. 2. Remove all of Wash Buffer B from mRNA-Dynabead tube. 3. Add 10 μl of RT mix to each sample. 4. Mix gently by flicking. 5. Centrifuge briefly. Repeat mixing and spin. 6. Incubate for 1–2 h at 42°C rotating in hybridization oven.

4.5. Agarose Bead Embedding of DNA

1. Remove Protease K-treated DNA tubes from waterbath. Place in heating block at >95°C. 2. Add 24 μl 2% LMP agarose (pre-warmed at >95°C) under mineral oil. 3. Mix gently by pipetting. Ensure the bead is well mixed.

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4. Incubate for 3 min >95°C to inactivate the Protease K. 5. Incubate for 10 min on ice to allow agarose bead to harden. 4.6. Denaturation of DNA

1. Remove oil from chilled, hardened agarose bead. 2. Add 1 ml 0.1 M NaOH to each tube. Invert 5–6 times (see Note 6). 3. Incubate for 15 min at 37°C in a waterbath, inverting every 3–4 min.

4.7. Bisulfite Treatment of DNA

1. Spin gently (